Fiber cells of cotton (Gossypium hirsutum L. and other Gossypium species including G. barbadense L., G. arboreum L., and G. herbaceous L.), a crop of enormous economic importance to world-wide agriculture, are differentiated epidermal cells of the seed coat. At maturity, the fiber cell, considered from inside to outside, consists of a cell lumen, secondary cell wall, primary cell wall, and thin waxy cuticle. The primary cell wall is made up of pectic compounds, hemicellulose components, cellulose, and protein. The secondary cell wall consists mainly (about 95%) of cellulose with small percentages of other components not yet conclusively identified.
Cotton fiber development is characterized by the stages of initiation, primary cell wall deposition, secondary cell wall deposition, and dessication. During the primary wall stage of fiber development, primary wall deposition occurs to facilitate fiber elongation. During the secondary wall stage of fiber development, secondary wall deposition occurs to accomplish fiber thickening. Primary and secondary wall deposition involve the synthesis of all the cell wall components characteristic of each stage and the assembly of the molecules into an organized cell wall outside the plasma membrane. Many hundred of genes are required for the differentiation and development of plant fiber. Work on in vitro translated fiber proteins (Delmer et al., “New Approaches to the Study of Cellulose Biosynthesis,” J. Cell Sci. Suppl., 2:33–50 (1985)), protein isolated from fiber (Graves and Stewart, “Analysis of the Protein Constituency of Developing Cotton Fibers,” J. Exp. Bot., 39:59–69 (1988)), and analysis of particular genes (Wilkins et al., “Molecular Genetics of Developing Cotton Fibers,” in Basra, ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing, Haworth Press:New York, p. 231–270 (1999)) clearly suggests differential gene expression during various developmental stages of the cell. However, only a few of the genes involved in the biosynthesis of the large numbers of fiber-specific or fiber-enhanced structural proteins, enzymes, polysaccharides, or waxes have been identified (John et al., “Gene Expression in Cotton (Gossypium hirsutum L.) Fiber: Cloning of the mRNAs,” Proc. Natl. Acad. Sci. USA, 89:5769–5773 (1992); John, “Characterization of a Cotton (Gossypium hirsutum L.) Fiber mRNA (Fb-B6),” Plant Physiol., 107:1477–1478 (1995)). Since these genes and their interactions with environment determine the quality of fiber, their identification and characterization is considered to be an important aspect of cotton crop improvement.
In particular, how secondary cell walls are synthesized, how they aid plant function, adaptation, and defense, and how their properties translate into industrial utility are important questions related to basic biological mechanisms, ecology, and plant improvement. Plant secondary cell walls are synthesized in some specialized cell types to facilitate particular functions, such as long-range conduction of water (tracheary elements), control of transpiration (guard cells), and dispersal of seeds (cotton fibers). These secondary cell walls have a higher content of high tensile strength cellulose, usually exceeding 40% by weight, and are much thicker than primary cell walls. Consequently, their hemicellulose, pectin, and protein content is reduced, with the most extreme reduction occurring in the case of cotton fiber secondary cell walls which are about 95% cellulose. On the other hand, during primary wall deposition, the cellulose content is typically 9–30% (w/w) (Meinert et al., “Changes in Biochemical Composition of the Cell Wall of the Cotton Fiber During Development,” Plant Physiol., 59:1088–1097 (1977); Darvill et al., “The Primary Cell Walls of Flowering Plants,” The Biochemistry of Plants, 1:91–162 (1980); Smook, Handbook for Pulp and Paper Technologists, Vancouver, Canada:Angus Wilde Publications, p. 15 (1992)). Because secondary cell walls are strong and represent a bulk source of chemical cellulose, they have been exploited as important renewable resources, for example in wood and cotton fibers.
Cotton fiber cells have two distinct developmental stages that include secondary wall deposition. Many studies of fiber length and weight increase, morphology, cell wall composition, cellulose synthesis rates, and gene expression have confirmed the summary of the stages of fiber development presented below, and recent reviews contain many primary references to confirm these facts (Delmer, “Cellulose Biosynthesis in Developing Cotton Fibers,” in Basra, ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing,” New York, N.Y.:Haworth Press, pp. 85–112 (1999); Ryser, “Cotton Fiber Initiation and Histodifferentiation,” in Basra ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing,” New York, N.Y.:Haworth Press, pp. 1–46 (1999); Wilkins et al., “Molecular Genetics of Developing Cotton Fibers, in Basra, ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing,” New York, N.Y.:Haworth Press, pp. 231–270 (1999)). Following fiber initiation by bulging of an epidermal cell above the surface, fiber elongation begins. Primary wall deposition is required to facilitate fiber elongation, and primary wall deposition continues alone for at least 12 days after fiber initiation. This period represents exclusively the primary wall stage of fiber development. Then, secondary wall deposition begins while primary wall deposition continues, albeit usually at a slower rate. This stage of fiber development represents the transition between primary and secondary wall deposition, and it typically begins in G. hirsutum L. between 14–17 days post anthesis (DPA). Subsequently, fiber elongation and primary wall deposition cease, typically between 18–24 DPA, and secondary wall deposition persists exclusively until 34–50 DPA. Variation in the time of initiation and duration of each phase of fiber development depends on the cotton cultivar and the temperature conditions (DeLanghe, “Lint Development” in Mauney, eds., Cotton Physiology, pp. 325–349, The Cotton Foundation, Memphis, Tennessee (1986); Haigler et al., “Cultured Cotton Ovules as Models for Cotton Fiber Development Under Low Temperatures,” Plant Physiol., 95:88–96 (1991); Thaker et al., “Genotypic Variation and Influence of Diurnal Temperature on Cotton Fiber Development,” Field Crops Research, 22:129–141 (1989)). For example, in field-grown G. barbadense L., extensive secondary wall deposition did not occur until after 20 DPA, elongation continued until 39 DPA, and secondary wall deposition ceased at 48 DPA (Schubert et al., “Growth and Development of the Lint Fibers of Pima S-4 Cotton,” Crop Sci., 16:539–543 (1976)).
The rates of cellulose synthesis change from low, to medium, to high, respectively, at the primary wall, transition, and secondary wall stages of fiber development (Meinert et al., “Changes in Biochemical Composition of the Cell Wall of the Cotton Fiber During Development,” Plant Physiol., 59:1088–1097 (1977); Martin, “Cool-Temperature-Induced Changes in Metabolism Related to Cellulose Synthesis in Cotton Fibers, Ph.D. dissertation, Texas Tech University, Lubbock, Tex., U.S.A. (1999)). An example is found in fibers developing on cultured cotton ovules at an optimum temperature of constant 34° C., which maximizes the rate of progression through the stages of fiber development. Combining results from fibers on ovules of two cotton cultivars of G. hirsutum L. cultured under this condition, primary wall deposition along with a low rate of cellulose synthesis occurs until 12–14 DPA, the transition between primary and secondary wall deposition and an intermediate rate of cellulose synthesis begins at 14–16 DPA, and secondary wall deposition continues along with a high rate of cellulose synthesis beginning at 16–21 DPA (Martin, “Cool-Temperature-Induced Changes in Metabolism Related to Cellulose Synthesis in Cotton Fibers, Ph.D. dissertation, Texas Tech University, Lubbock, Tex. U.S.A. (1999)). Other biochemical features demonstrate that the initiation of secondary wall deposition via an intermediate rate of cellulose synthesis at the transition stage marks a distinct developmental event: the cellulose content of new wall material rises sharply so that the overall percentage of cellulose in the whole fiber wall doubles in one day (Meinert et al., “Changes in Biochemical Composition of the Cell Wall of the Cotton Fiber During Development,” Plant Physiol., 59:1088–1097 (1977)), the respiration rate transiently declines, and the intercellular pools in the fiber of UDP-glucose and glucose-6-P begin to rise (Martin, “Cool-Temperature-Induced Changes in Metabolism Related to Cellulose Synthesis in Cotton Fibers, Ph.D. dissertation, Texas Tech University, Lubbock, Tex. U.S.A. (1999)). These are signs of an abrupt onset of secondary wall deposition (DeLanghe, “Lint Development” in Mauney, eds., Cotton Physiology, pp. 325–349, The Cotton Foundation, Memphis, Tenn. (1986)).
Primary and secondary wall deposition appear to be controlled by different genetic factors (Kohel et al., “Fiber Elongation and Dry Weight Changes in Mutant Lines of Cotton,” Crop Sci., 14:471–474 (1974)), and both stages can likely be manipulated independently by genetic engineering to achieve the longer, stronger, finer (smaller diameter), and more mature (as related to the secondary wall thickness) fiber that the textile industry desires. However, this goal can only be achieved by knowing more about the genes that control and contribute to each stage of fiber development.
After the fibers mature, the protective boll opens and the dried fiber often hangs for several weeks on the plant until the whole crop matures (unless stopped by killing cold temperatures) and vegetative growth dies or is killed chemically to allow harvest. During this period, fibers are subject to degradation by enzymatic activity of fungi, which is often enhanced by wet fall weather (Simpson et al., “The Geographical Distribution of Certain Pre-Harvest Microbial Infections of Cotton Fiber in the U.S. Cotton Belt,” Plant Disease Reporter, 55:714–718 (1971)). In some years, this field-waiting time causes substantial deterioration of the grade of the fiber so that the producer receives a discounted price and the production of quality yams and fabrics is jeopardized. Therefore, cotton production efficiency will be improved by more knowledge of how to bring the fibers undamaged from the field to textile plant. Relevant to achieving this goal is a better understanding of endogenous protections against fungal degradation that could be introduced or enhanced in the fiber.
Particularly because of their defensive role in plants (Gooday, “Aggressive and Defensive Roles for Chitinases,” in Jolles, eds., Chitin and Chitinases, Birkhäuser Verlag:Basel, pp. 157–170 (1999)), numerous chitinase genes and proteins have been characterized in diverse plant species. The chitinase gene and protein family has been the subject of many reviews (including Graham et al., “Cellular Coordination of Molecular Responses in Plant Defense,” Molecular Plant-Microbe Interactions: MPMI, 4:415–422 (1991); Cutt et al., “Pathogenesis-Related Proteins,” in Boller, eds., Genes Involved in Plant Defense, Springer Verlag/N.Y. pp. 209–243 (1992); Meins et al., “The Primary Structure of Plant Pathogenesis-Related Glucanohydrolases and Their Genes,” in Boller, eds., Genes Involved in Plant Defense, Springer Verlag/N.Y., p. 245–282 (1992); Collinge et al., “Plant Chitinases,” Plant J., 3:31–40 (1993); Sahai et al., “Chitinases of Fungi and Plants: Their Involvement in Morphogenesis and Host-Parasite Interaction,” FEMS Microbiology Rev., 11:317–338 (1993); Meins et al., “Plant Chitinase Genes,” Plant Molecular Biology Reporter, 12:522–528 (1994); Hamel et al., “Structural and Evolutionary Relationships Among Chitinases of Flowering Plants,” Journal of Molecular Evolution, 44:614–624 (1997)) and of an edited book (Jolles, eds. Chitin and Chitinases, Birkäuser Verlag:Basel, 340 pp (1999)). These sources contain many primary references to the well known facts summarized below. Chitinases are among a group of genes that are inducible in plants by pathogen attack, corresponding to the frequent occurrence of chitin in fungal cell walls and insect exoskeletons. In their defensive role, chitinases catalyze the hydrolysis of chitin. Structural chitin occurs as crystalline microfibrils composed of a linear homopolymer of β-1,4-linked N-acetyl-D-glucosamine residues, (GlcNAc)n. Chitin hydrolysis defends the plant against predators or pathogens, particularly invading fungi, by weakening or dissolving their body structure. Especially in combination with β-1,3-glucanases that serve to uncoat the chitin microfibrils, chitinases can inhibit the growth of many fungi by causing hyphal tip lysis due to a weakened hyphal wall. This has been shown by inhibition of fungal growth in cultures as well as in transgenic plants that exhibit reduced pathogen damage in correlation with increased chitinase activity. Gene expression or activity of chitinases with a probable defensive function have previously been characterized in cotton leaves and roots (Liu et al., “Detection of Pathogenesis-Related Proteins in Cotton Plants,” Physiological and Molecular Plant Pathology, 47:357–363 (1995); Hudspeth et al., “Characterization and Expression of Chitinase and 1,3-β-Glucanase Genes in Cotton,” Plant Molecular Biology, 31:911–916 (1996); Dubery et al., “Induced Defence Responses in Cotton Leaf Disks by Elicitors From Verticillium dahliae,” Phytochemistry, 44: 1429–1434 (1997)).
Some chitinases are induced upon fungal invasion, accumulating around invading fungal hyphae (Benhamou et al., “Subcellular Localization of Chitinase and Its Potential Substrate in Tomato Root Tissues Infected With Fusarium Oxysporium F.Sp. Radicislycopersici,” Plant Physiology, 92:1108–1120 (1990); Wubben et al., “Subcellular Localization of Plant Chitinases and 1,3-β-Glucanases in Cladosporium Fulvum (Syn. Fulvia Fulva)-Infected Tomato Leaves,” Physiological and Molecular Plant Pathology 41:23–32 (1992)). Other chitinases apparently occur constitutively in plant parts that are particularly susceptible to invasion such as epidermal cells, root cortical cells, stomates, flower parts, and vascular cells. These conclusions arise from both localization of chitinase mRNA by in situ hybridization and analysis of patterns of expression of the GUS reporter gene under control of chitinase promoters (Samac et al., “Developmental and Pathogen-Induced Activation of the Arabidopsis Acidic Chitinase Promoter,” The Plant Cell, 3:1063–1072 (1991); Zhu et al., “Stress Induction and Developmental Regulation of a Rice Chitinase Promoter in Transgenic Tobacco,” The Plant Journal, 3:203–212 (1993); Büchter et al., “Primary Structure and Expression of Acidic (Class II) Chitinase in Potato,” Plant Molecular Biology, 35:749–761 (1997); Ancillo et al., “A Distinct Member of the Basic (Class I) Chitinase Gene Family in Potato is Specifically Expressed in Epidermal Cells,”. Plant Molecular Biology, 39:1137–1151 (1999)). In another case of possible anticipation of fungal invasion in disrupted tissues, ethylene induces chitinase in bean abscission zones (del Campillo et al., “Identification and Kinetics of Accumulation of Proteins Induced by Ethylene in Bean Abscission Zones,” Plant Physiology, 98:955–961 (1991)). None of these studies included analysis of cotton fibers or showed the presence of chitinase activity or chitinase-related proteins in cotton fibers.
Other studies show that some chitinases have a developmental role in plants, although authentic structural chitin is not a natural part of the plant body (Meins et al., “The Primary Structure of Plant Pathogenesis-Related Glucanohydrolases and Their Genes,” In Boller, eds., Genes Involved in Plant Defense, Springer Verlag/N.Y., p. 245–282 (1992)). It has been shown that chitinase isoforms with developmental roles are at least sometimes distinct from those with roles in stress responses and defense (Mauch et al., “Antifungal Hydrolases in Pea Tissue. I. Purification and Characterization of Two Chitinases and Two β-1,3-Glucanases Differentially Regulated During Development and in Response to Fungal Infection,” Plant Physiology, 87:325–333 (1988)). Defensive chitinases bind to short stretches (probably 3–6 residues) of a single N-acetyl-glucosamine chain prior to cleaving the inter-sugar bond (Robertus et al., “The Structure and Action of Chitinases,” in Jolles, eds., Chitin and Chitinases, Birkäuser Verlag:Basel, pp. 125–136 (1999)). Therefore, enzymes in the chitinase family can also bind to oligomers of N-acetyl-glucosamine within other molecules such as glycoproteins or signalling molecules. Such molecules may have roles in signal transduction to regulate gene expression cascades required for developmental transitions or in the biosynthetic processes that implement the developmental program.
Previous research on the regulation of secondary wall deposition or function at the molecular level focused on a few genes involved in cellulose, hemicellulose, lignin, or protein biosynthesis. Among these pathways, lignin synthesis has been most fully explored and manipulated in transgenic plants (Merkle et al., “Forest Biotechnology,” Current Opinion in Biotechnology, 11:298–302 (2000)). However, cotton fibers contain no lignin (Haigler, “The Functions and Biogenesis of Native Cellulose,” in Zeronian, eds., Cellulose Chemistry and Its Applications, Ellis Horwood:Chichester, England, pp. 30–83 (1985)). Hemicellulose polysaccharides within some secondary walls include xylans and glucomannans, but cotton fiber secondary walls do not contain significant quantities of any similar molecule (Haigler, “The Functions and Biogenesis of Native Cellulose,” in Zeronian, eds., Cellulose Chemistry and Its Applications, Ellis Horwood:Chichester, England, pp. 30–83 (1985)). Only two proteins with possible structural roles in the cotton fiber secondary wall have been identified. One of these, H6, is an arabinogalactan-type protein that accumulates to detectable levels during secondary wall deposition, although the expression of its gene begins during rapid elongation including primary wall deposition (John et al., “Characterization of mRNA for a Proline-Rich Protein of Cotton Fiber,” Plant Physiology, 108:669–676 (1995)). The second, FbL2A, lacks homology to any known protein, but its highly repetitive sequence and high hydrophilicity suggest that it may have a structural role or protect cotton fibers during dessication. The expression of its gene begins weakly at the primary to secondary wall transition (15 DPA) and is stronger by 20 DPA (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)).
Enzymes that increase in gene expression and/or activity during cotton fiber secondary wall deposition and that relate to the regulation of cellulose synthesis include cellulose synthase, sucrose synthase, sucrose phosphate synthase, and UDP-glucose pyrophosphorylase (Basra et al., “Sucrose Hydrolysis in Relation to Development of Cotton (Gossypium spp.) Fibres,” Indian Journal of Experimental Botany, 28:985–988 (1990); Wäfler et al., “Enzyme Activities in Developing Cotton Fibres,” Plant Physiology and Biochemistry, 32:697–702 (1994); Amor et al., “A Membrane-Associated Form of Sucrose Synthase and its Potential Role in Synthesis of Cellulose and Callose in Plants,” Proc. Nat'l. Acad. Sci. U.S.A., 92:9353–9357 (1995); Pear et al., “Higher Plants Contain Homologs of the Bacterial celA Genes Encoding the Catalytic Subunit of Cellulose Synthase,” Proc. Nat'l. Acad. Sci. U.S.A., 93:12637–12642 (1996); Tummala, “Response of Sucrose Phosphate Synthase Activity to Cool Temperatures in Cotton,” M.S. thesis, Texas Tech University, Lubbock, Tex. (1996)). UDP-glucose pyrophosphorylase converts glucose-1-P to UDP-glucose or mediates the reverse reaction. In the context of cellulose synthesis, sucrose synthase is thought to degrade sucrose and supply UDP-glucose to cellulose synthase. Cellulose synthase transfers the glucose to the elongating β-1,4-linked cellulose polymer while free UDP is recycled to sucrose synthase. Sucrose phosphate synthase may use fructose-6-P (e.g. that derived from the fructose released by the degradative action of sucrose synthase) and UDP-glucose to synthesize additional sucrose to support cellulose synthesis (Delmer, “Cellulose Biosynthesis in Developing Cotton Fibers,” in Basra, ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing, Haworth Press:New York, pp. 85–112 (1999)).
All of the enzymes just discussed operate within the pathways of sugar metabolism leading to the formation of the β-1,4-linked glucan polymer. Evidence from other systems and cotton fibers implicates other possible points of regulation of cellulose synthesis coincident with or after formation of the glucan polymer, although the relevant pathways and proteins are incompletely understood. Examples of other relevant proteins include β-1,4-glucanase that may act as a glucan chain editor or in some other role (Delmer, “Cellulose Biosynthesis: Exciting Times For a Difficult Field of Study,” Ann. Rev. Plant Physiol. Mol. Biol., 50:245–276 (1999)) and glycoproteins that may act as primers for cellulose biosynthesis (Lukowitz et al., “Arabidopsis cyt1 Mutants are Deficient in a Mannose-1-Phosphate Guanyltransferase and Point to a Requirement of N-Linked Glycosylation for Cellulose Biosynthesis,” Proc. Nat'l. Acad. Sci. U.S.A., 98:2262–2267 (2001)). Mutations that down-regulate β-1,4-glucanase or glycoprotein synthesis cause reduced cellulose content in other systems (Nicol et al., “Plant Cell Expansion: Scaling the Wall,” Current Opinion in Cell Biology, 1:12–17 (1998); Lukowitz et al., “Arabidopsis cyt1 Mutants are Deficient in a Mannose-1-Phosphate Guanyltransferase and Point to a Requirement of N-Linked Glycosylation for Cellulose Biosynthesis,” Proc. Nat'l. Acad. Sci. U.S.A., 98:2262–2267 (2001)). In Arabidopsis, mutation of xylem secondary wall specific cellulose synthase genes causes reduced cellulose content and weak xylem walls (Turner et al., “Collapsed Xylem Phenotype of Arabidopsis Identifies Mutants Deficient in Cellulose Deposition in the Secondary Cell Wall,” Plant Cell, 9:689–701 (1997)). In cotton, increased expression of spinach sucrose phosphate synthase causes increased cellulose content in fiber walls of plants growing under a cool night cycle (Haigler et al., “Transgenic Cotton Over-Expressing Sucrose Phosphate Synthase Produces Higher Quality Fibers With Increased Cellulose Content and Has Enhanced Seed Cotton Yield,” Abstract 477. In: Proceedings of Plant Biology 2000, July 15–19, San Diego, Calif., American Society of Plant Physiologists, Rockville, Md., (2000)). These findings indicate that it is possible to manipulate the quantity of cellulose in secondary walls, but presently there is insufficient identification of target genes that might be beneficially manipulated.
Studies that identify genes that are under tissue-specific and developmental regulation are important in understanding the roles of proteins in fiber development and cell-wall architecture (John, “Structural Characterization of Genes Corresponding to Cotton Fiber mRNA, E6: Reduced E6 Protein in Transgenic Plants by Antisense Gene,” Plant Mol. Biol., 30:297–306 (1996)). In addition, such genes and their regulatory elements are important tools for fiber modification through genetic engineering (John, “Prospects for Modification of Fibers Through Genetic Engineering of Cotton,” in Gebelein, eds., Industrial Biotechnological Polymers, Lancaster, Pa.:Technomic, pp. 69–79 (1995); John, “Genetic Engineering Strategies for Cotton Fiber Modification. In: A. S. Basra (ed.), Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing, The Haworth Press, New York, pp. 271–289 (1999)).
In many instances, it would be desirable for a transgene to be developmentally regulated to have exclusive or preferential expression in fiber cells at a proper developmental stage. This regulation can be most expeditiously accomplished by a promoter capable of preferential promotion.
Promoters are DNA elements that direct the transcription of RNA in cells. Together with other regulatory elements that specify tissue and temporal specificity of gene expression, promoters control the development of organisms. Thus, there has been a concerted effort in identifying and isolating promoters from a wide variety of plants and animals.
Many promoters function properly in heterologous systems. For example, promoters taken from plant genes such as rbcS, Cab, chalcone synthase, and protease inhibitor from tobacco and Arabidopsis are functional in heterologous transgenic plants. (Benfey et al., “Regulated Genes in Transgenic Plants,” Science, 244:174–181, (1989)). Specific examples of transgenic plants include tissue-specific and developmentally regulated expression of soybean 7s seed storage protein gene in transgenic tobacco plants (Chen et al., “A DNA Sequence Element That Confers Seed-Specific Enhancement to a Constitutive Promoter,” EMBO J., 7:297–302, (1988)) and light-dependent organ-specific expression of Arabidopsis thaliana chlorophyll a/b binding protein gene promoter in transgenic tobacco (Ha et al., “Identification of Upstream Regulatory Elements Involved in the Developmental Expression of the Arabidopsis thaliana Cab-1 Gene,” Proc. Natl. Acad. Sci. USA, 85:8017–8021, (1988)). Similarly, anaerobically inducible maize sucrose synthase-1 promoter activity was demonstrated in transgenic tobacco (Yang et al., “Maize Sucrose Synthase-1 Promoter Directs Phloem Cell-Specific Expression of Gus Gene in Transgenic Tobacco Plants,” Proc. Natl. Acad. Sci. USA, 87: 4144–4148, (1990)). Tomato pollen promoters were found to direct tissue-specific and developmentally regulated gene expression in transgenic Arabidopsis and tobacco (Twell et al., “Pollen-Specific Gene Expression in Transgenic Plants Coordinate Regulation of Two Different Tomato Gene Promoters During Microsporogenesis,” Development, 109:705–714, (1990)). Thus, some plant promoters can be utilized to express foreign proteins in plant tissues in a developmentally regulated fashion.
Tissue-specific and developmentally regulated expression of genes has also been shown in fiber cells. Several of these, such as E6, vacuolar ATPase, and lipid transfer-type proteins, have strong expression in cotton fibers during primary wall deposition (Wilkins et al., “Molecular Genetics of Developing Cotton Fibers,” in Basra, ed., Cotton Fibers: Developmental Biology, Quality Improvement, and Textile Processing, Haworth Press:New York, p. 231–270 (1999); Orford et al., “Expression of a Lipid Transfer Protein Gene Family During Cotton Fibre Development,” Biochimica and Biophysica Acta, 1483:275–284 (2000)). Other genes show transient expression during fiber development. For example, Rae is transiently expressed at the primary to secondary wall stage transition (Delmer et al., “Genes Encoding Small GTP-Binding Proteins Analogous to Mammalian Rae are Preferentially Expressed in Developing Cotton Fibers,” Mol. Gen. Genet., 248:43–51 (1995)) and another lipid transfer-type protein, FS18A (Orford et al., “Characterization of a Cotton Gene Expressed Late in Fibre Cell Elongation,” Theoretical and Applied Genetics, 98:757–764 (1999)), is transiently expressed at 24 DPA during secondary wall deposition. Another gene, H6, is expressed between 10–24 DPA, which includes both primary and early secondary wall deposition, but, after the promoter of this gene has commenced activity during primary wall deposition, there is post-transcriptional control of gene expression so that H6 protein accumulates only during secondary wall deposition at 15–40 DPA (John et al., “Characterization of mRNA for a Proline-Rich Protein of Cotton Fiber,” Plant Physiology, 108:669–676 (1995)). At the level of gene expression, only certain cellulose synthase genes have been previously shown to have preferential and prolonged expression in cotton fibers during secondary wall deposition, although there were lower levels of expression during primary wall deposition and in other parts of the plant. In addition, cellulose synthase did not show strong expression until 20 DPA, with only weak expression observed at 17 DPA (Pear et al., “Higher Plants Contain Homologs of the Bacterial celA Genes Encoding the Catalytic Subunit of Cellulose Synthase,” Proc. Nat'l. Acad. Sci. U.S.A., 93:12637–12642 (1996)). Another gene, FbL2A, is up-regulated at the primary to secondary wall transition (weakly at 15 DPA and strongly at 20 DPA) (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)).
Any of the genes expressed in cotton fibers are candidates for containing useful promoters, with a particular class of useful promoter being those that are preferentially expressed in fibers and/or secondary walled cells compared to other cell types and/or developmental stages. Promoters expressing preferentially during the primary wall stage of fiber development will also have particular uses. It may also be advantageous to have promoters that express in cotton fibers and certain restricted classes of cells (for example, secondary walled cells) in other parts of the cotton plant. Promoters with different strength are also valuable because different genetic engineering goals may be best accomplished with different amounts of the foreign protein being present in the cell or tissue. The biotechnology industry dealing with any particular species will ultimately desire and need a “toolbox” of promoters so that the most appropriate one for any particular use may be chosen. Each promoter will have a combination of unique characteristics in terms of cell, tissue, or developmental specificity of driving gene expression, strength of gene expression, degree of susceptibility to positional effects of gene insertion on level of gene expression, susceptibility to gene silencing, and any number of other similar phenomena that affect transcription.
A few promoters of genes expressed in cotton fibers have been isolated and tested in stably transformed cotton and with relationship to two or more time points in the time course of fiber development, where the testing involved promoter fusions with “reporter” genes or genes that are part of putatively useful genetic engineering strategies. Three major patterns were observed, one typified by the Gh10 promoter (for an acyl carrier protein), which drives foreign gene expression throughout fiber development (Song et al., “Expression of a Promoter from a Fiber-Specific Acyl Carrier Protein Gene in Transgenic Cotton Plants,” Proc. Beltwide Cotton Conf., 1:486–488 (1998)). A second pattern was typified by the E6 promoter, which drives gene expression preferentially at the primary wall stage of cotton fiber development (U.S. Pat. No. 5,521,078 to John). A third pattern was shown by the promoter of the FbL2A gene. This promoter was tested by fusion to two reporter genes (polyhydroxyalkanoic acid synthase or PHA synthase, an enzyme that was detected with a specific antibody, and acetoacetyl-CoA reductase, an enzyme with activity that was monitored by enzyme assay) and transformation of cotton (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)). However, by use of the two promoter/reporter gene constructs, somewhat contradictory data were provided on the utility of the promoter of FbL2A. When the acetoacetyl-CoA reductase gene was under control of the FbL2A promoter, acetoacetyl-CoA reductase activity was detected in cotton fibers during primary wall deposition at 5–10 DPA, and there was consistent, substantial activity by 20 DPA during secondary wall deposition, peak activity at 35 DPA, and continued activity until 45 DPA among a family of independently transformed plants (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)). However, the enzyme activity after 20 DPA could be due to long-lived messenger RNA or protein synthesized at 15–20 DPA, which is the period when the data directly show FbL2A gene expression under control of its own promoter. In contrast, when the PHA synthase gene was under the control of the FbL2A promoter, Western blotting to detect PHA synthase immunologically in a transformed plant showed only very weak signal during secondary wall deposition at 20 DPA, a trace signal at 25 DPA, and no signal at 30 or 35 DPA (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)). Furthermore, the PHA synthase gene under the control of the putatively consitutive 35S promoter from CaMV virus correlated with detection of PHA synthase protein strongly during primary wall deposition at 10 DPA and weakly continuing through 35 DPA of secondary wall deposition. The comparative patterns during secondary wall deposition are consistent with transient expression of the FbL2A promoter around 20 DPA of secondary wall deposition. The data also show that the FbL2A promoter will drive weak foreign gene expression in cotton fibers during primary wall deposition (Rinehart et al., “Tissue-Specific and Developmental Regulation of Cotton Gene FbL2A,” Plant Physiology, 112:1331–1341 (1996)). Further, graphical data in U.S. Pat. No. 6,211,430 to John showed that the FbL2A promoter activity was not substantial until 20 DPA (about 5 days past the onset of secondary wall deposition) and that 50% of maximal foreign acetoacetyl-CoA reductase activity was not detectable until 31 DPA (about 16 days past onset of secondary wall deposition).
Thus, it would be useful to have a promoter that would drive gene expression preferentially and strongly in secondary walled cells—fibers, in particular—throughout secondary wall deposition, i.e., strongly and continuously (e.g. at ≧50% of its maximal activity) from the initiation of secondary wall deposition to its termination. The initiation of secondary wall deposition is defined as the time when the dry weight/unit length of a cotton fiber begins to increase or when the dry weight/unit surface area of any cell begins to increase via synthesis of new wall material containing more than 30% (w/w) of cellulose. In the case of cotton fiber of G. hirsutum L., this is expected to occur between 14–17 DPA when cotton plants are grown under typical conditions in the greenhouse or the field (day temperature of 26–34° C., night temperature of 20–26° C., light intensity greater than or equal to 1000 μeinsteins/m2/s, with adequate water and mineral nutrition). Furthermore, it would be useful to have a promoter that would drive gene expression only or preferentially in secondary walled cells such as fibers while excluding or minimizing expression in other cell types.
The present invention is directed to achieving these objectives.